生物化学

Metamorphosis in class Insecta is defined as the drastic transition of an organism from a larval stage to a morphologically very different adult form. Insects such as Drosophila melanogaster undergo metamorphosis, which requires precise temporal control, exerted by ecdysteroids, the primary regulators of these developmental transitions. The active form of ecdysone is 20-hydroxyecdysone (20E), which acts in pulses that trigger the onset of different developmental stages. A massive pulse of 20E occurs between the prepupal and pupal stages, triggering pupariation. Precise quantification of 20E levels is critical for understanding how this hormonal signaling governs the developmental timing of different stages. This protocol is sensitive for detecting low levels of 20E and describes a methanol-based extraction method coupled with a competitive ELISA to quantify whole-body 20E levels in third-instar Drosophila larvae. Further, this protocol enables detection of differences in ecdysone titers among genotypes affecting ecdysone biosynthesis, particularly at the late third-instar larval stage. This protocol can also be utilized for disease-causing insects (Aedes aegypti), as well as economically relevant beneficial insects (Apis mellifera), to study the effects of ecdysteroid analogs during development.

Understanding gene expression within defined neuronal populations is essential for dissecting the cellular and molecular diversity of the brain. mRNA assays provide a direct readout of gene expression, capturing transcriptional changes that may precede or occur independently of protein abundance, whereas protein assays reflect the cumulative effects of translation, modification, and degradation. Moreover, in histological analysis, immunohistochemical protein detection results in visually diffuse labeling, which makes it difficult to quantitatively assess levels and locations of expression at high resolution. Here, we present a protocol that allows for mRNA detection in single neuronal cell types with a high degree of sensitivity and anatomical resolution. This protocol combines fluorescent in situ hybridization (FISH) with immunohistochemistry (IHC) on the same tissue section. Briefly, FISH is carried out by ACDBio RNAscope^® fluorescent in situ hybridization technology, which involves processing the tissue sections, followed by signal amplification. This involves target retrieval, probe hybridization, and signal enhancement. Then, the tissue section is processed for IHC, which involves blocking nonspecific sites and incubation with primary antibodies, followed by development of a fluorescent signal with secondary antibodies. Typically, visual mRNA detection with FISH can be seen as individual puncta, whereas targeting the protein with an antibody results in filled cells or processes. The variation in staining pattern allows for the quantification of distinct mRNA transcripts within different neuronal populations, which renders co-localization analyses easy and efficient.

Fluorescence in situ hybridization (FISH) can be employed to study the expression and subcellular localization of nucleic acids by using labeled antisense strands that hybridize with the target RNA or DNA molecules. Likewise, immunofluorescence antibody staining (IF) takes advantage of the specific interaction between a fluorophore-labeled antibody and its corresponding antigen. This protocol reports the combination of RNA-FISH and IF antibody staining for simultaneous detection of both RNA transcripts and proteins of interest in routine formalin-fixed paraffin-embedded (FFPE) bone marrow biopsy samples. Herein, we provide a detailed description of the methodology that we have developed and optimized to study the spatial expression of two transcripts—TGFB1 and PDGFA1—in human hematopoietic (CD45⁺) and non-hematopoietic (CD271⁺) cells in the bone marrow of patients with acute lymphoblastic leukemia (ALL).

Three-dimensional immunohistochemistry (3D-IHC) shows the organization of molecular assemblies in the context of tissue architecture. Deep and rapid antibody penetration into 3D tissues and highly sensitive detection are crucial for high-throughput analysis of 3D-IHC imaging. Here, we provide a detailed protocol for a nanobody (nAb)-based 3D-IHC technique, namely POD-nAb/FT-GO 3D-IHC, for high-speed and high-sensitivity detection of targets within 1-mm-thick mouse brain tissues. Peroxidase-fused nAb (POD-nAb) is a genetically encoded recombinant antibody, which consists of a camelid nAb and a variant of horseradish peroxidase, and fluorochromized tyramide-glucose oxidase (FT-GO) is a fluorescent tyramide signal amplification (TSA) system. POD-nAb/FT-GO 3D-IHC incorporates three main components: 1) tissue permeabilization, 2) POD-nAb binding, and 3) 3D-TSA reaction with FT-GO. POD-nAbs enhance signal penetration depth and allow for highly sensitive detection when combined with FT-GO signal amplification. By using the 3D-IHC protocol provided herein, we can visualize target molecules in mouse brain tissues of 1-mm thickness with drastic signal enhancement within three days. This protocol for POD-nAb/FT-GO 3D-IHC could facilitate structural and molecular interrogation of 3D tissues.

Small ubiquitin-related modifiers (SUMOs) are covalently conjugated onto the proteome and serve as signaling molecules in many aspects of eukaryotic cell biology, from S. cerevisiae and C. elegans to H. sapiens. The conjugatable SUMO variants, SUMO1 and the almost identical SUMO2 and SUMO3 (designated SUMO2/3), are processed by an E1(SAE1:SAE2)-E2(UBC9)-E3 enzyme cascade to produce SUMO-modified proteins. The prerogative of the SUMO biology field is to identify and study the specific proteins undergoing SUMOylation, which grants us insights into the biological pathway of interest. This protocol was developed using the human osteosarcoma cell line U2OS to enable the investigation of SUMO conjugates in mitosis, the cell division phase of the cell cycle. We enrich the cell population for mitotic cells, which are isolated and subjected to stringent lysis conditions involving a high concentration of SDS and DTT in RIPA buffer, to promote complete protein denaturation. The lysates in high SDS RIPA buffer are diluted to reduce the overall SDS concentration and undergo conventional immunoprecipitation using SUMO1- or SUMO2/3-specific antibodies bound to protein A/G agarose beads. The samples are then compatible with downstream readouts such as western blots and mass spectrometry. This protocol detects endogenous SUMOylated proteins and avoids exogenous SUMO overexpression, which can alter SUMO conjugate formation. Furthermore, this denaturing protocol ensures only SUMOylated proteins are immunoprecipitated, and not their interactors.

Cyclic GMP–AMP synthase (cGAS) is a key cytosolic double-stranded DNA sensor that activates innate immune responses. Upon binding double-stranded DNA, cGAS undergoes conformational activation and catalyzes the synthesis of the second messenger 2′3′-cyclic GMP–AMP (2′3′-cGAMP) from ATP and GTP. 2′3′-cGAMP then triggers a downstream signaling cascade that induces type-I interferon and inflammatory gene expression and has been shown to exert antitumor effects in the context of cancer. Accurate measurement of this enzymatic activity is therefore important for mechanistic studies. Traditional kinetic methods such as radiolabeling, HPLC, or mass spectrometry provide precise results but require specialized equipment and expertise. Here, we describe a rapid and accessible ELISA-based protocol to quantify 2′3′-cGAMP product formation and derive cGAS enzymatic parameters. Reactions are initiated with defined DNA ligands and quenched at multiple time points, and product accumulation is quantified by a commercially available 2′3′-cGAMP ELISA. Time course measurements are used to calculate initial velocities, which can be plotted against substrate concentration to obtain Michaelis–Menten parameters. This approach enables direct, product-specific quantification of 2′3′-cGAMP formation using only an absorbance plate reader. The protocol provides a sensitive and broadly applicable alternative to traditional methods, allowing laboratories without advanced instrumentation to perform reliable cGAS enzyme kinetics.

Time-lapse into immunofluorescence (TL into IF) imaging combines the wealth of information acquired during live-cell imaging with ease of access for static immunofluorescence markers. In the field of mechanobiology, connecting live and static imaging to visualize cell biology dynamics is often troublesome. For instance, nuclear blebs are deformations of the nucleus that often rupture spontaneously, leading to changes in the molecular composition of the nucleus and the nuclear bleb. Current techniques to connect cellular dynamics and their downstream effects via live-cell imaging, followed by immunofluorescence, often require third-party analysis programs or stage position measurements to accurately track cells. This protocol simplifies the connection between live and static imaging by utilizing a gridded imaging dish. In our protocol, cells are plated on a dish with an engraved coordinate plane. Individual cells are then matched from when the time-lapse ends to the immunofluorescence images simply by their known coordinate location. Overall, TL into IF offers a straightforward method for connecting dynamic live-cell with static immunofluorescence imaging, in an easy and accessible tool for cell biologists.

The antibody-uptake assay is a commonly used technique to monitor endocytosis of integral membrane proteins including transmembrane and glycosylphosphatidylinositol-anchored proteins (GPI-APs). The antibody-uptake assay typically involves incubating live cells with fluorophore-conjugated antibodies directed against the extracellular domain of the integral membrane protein of interest. Antibody uptake is then detected by flow cytometry or confocal microscopy. However, these detection modalities may be inaccessible to some labs or require extensive training to operate. Thus, we developed an easy and novel sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and western blot-based approach to the antibody-uptake assay that exploits the strong affinity between biotin and streptavidin. Instead of incubating cells with fluorophore-conjugated antibodies to monitor antibody uptake, our assay involves incubating cells with biotinylated antibodies, processing the cell lysates for western blot, and probing the membrane with detectably conjugated streptavidin. From preparation to quantification, this protocol requires less hands-on time than other approaches and is amenable to small-scale drug or siRNA screens. Here, we demonstrate the utility of our approach using the well-characterized misfolded GPI-AP, YFP-tagged C179A mutant of prion protein (YFP-PrP*), as our model substrate. YFP-PrP* constitutively traffics to the plasma membrane (PM), where it binds to anti-GFP antibody, and immediately undergoes endocytosis to lysosomes. To validate our protocol, we present measurements of antibody uptake under conditions known to enhance or inhibit YFP-PrP*’s traffic to the PM. Using this assay, we present new evidence that, under certain conditions, YFP-PrP* is able to undergo degradation via a pathway that does not involve exposure on the cell surface.

Nowadays, recombinant proteins are the focus of various research fields, and their use ranges from therapeutic investigations to cellular model systems for the development of therapeutic approaches. Cell systems used for the expression of recombinant proteins should be comparable in terms of yield and expression efficiency. In many research fields, it is desirable to obtain high protein concentrations. A method that combines an easy workflow with rapid results and affordable costs remains missing, and a standardized approach to determining protein concentration in transgenic cell lines is essential for more reliable data analysis. Our protocol demonstrates the cluster fluorescence-linked immunosorbent assay (FLISA), a technique that allows the exact quantification of comparable protein expression amounts. Moreover, it enables the detection of clustered or bound subunits of a protein without necessitating ultracentrifugation. In the present protocol, we demonstrate the utilization of two transgene cell lines, each expressing distinct recombinant proteins, to provide comparability of protein yields and detectable subunit clustering.

N-glycosylation is a ubiquitous post-translational modification (PTM) that regulates protein folding, stability, and biological function. Accurate identification and validation of N-glycosylation are therefore critical for understanding how glycosylation modulates protein activity. Here, we present a robust workflow for analyzing protein N-glycosylation in both animal and plant systems using peptide-N⁴-(N-acetyl-β-glucosaminyl) asparagine-amidase A and F (PNGase A and PNGase F). After enzymatic cleavage of the asparagine-linked N-glycans, samples are analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and western blotting (WB) to detect shifts in apparent molecular weight (MW) indicative of deglycosylation. Key steps include denaturing the protein to expose glycosylation sites, optimizing buffer conditions for PNGase F and A treatment, and comparing glycosylated vs. deglycosylated forms by electrophoretic mobility. A troubleshooting guide addresses common challenges, including incomplete deglycosylation and low transfer efficiency during WB, offering practical solutions to ensure reliable results. This protocol provides researchers with a standardized, cost-effective framework for investigating protein N-glycosylation in diverse systems, from cell lysates to purified proteins, in both animal and plant models.